Phosphorylation-Coupled Intramolecular Dynamics of

2222 Biophysical Journal Volume 104 May 2013 2222–2234

Phosphorylation-Coupled Intramolecular Dynamics of UnstructuredRegions in Chromatin Remodeler FACT

Manami Hashimoto,†††6 Noriyuki Kodera,{6 Yasuo Tsunaka,‡‡‡6 Masayuki Oda,** Mitsuru Tanimoto,k

Toshio Ando,{k Kosuke Morikawa,‡§* and Shin-ichi Tate†††*†Department of Mathematical and Life Sciences, Hiroshima University, Hiroshima, Japan; ‡Institute for Integrated Cell-Material Sciences(iCeMS), Kyoto University, Kyoto, Japan; §International Institute for Advances Studies, Kyoto, Japan; {Bio-AFM Frontier Research Center,College of Science and Engineering, Kanazawa University, Kanazawa, Japan; kSchool of Mathematics and Physics, College of Science andEngineering, Kanazawa University, Kanazawa, Japan; **Graduate School of Life and Environmental Sciences, Kyoto Prefectural University,Kyoto, Japan; ††SENTAN, Japan Science and Technology Agency, Tokyo, Japan; and ‡‡PRESTO, Japan Science and Technology Agency,Tokyo, Japan

ABSTRACT The intrinsically disordered region (IDR) of a protein is an important topic in molecular biology. The functional sig-nificance of IDRs typically involves gene-regulation processes and is closely related to posttranslational modifications such asphosphorylation. We previously reported that the Drosophila facilitates chromatin transcription (FACT) protein involved in chro-matin remodeling contains an acidic ID fragment (AID) whose phosphorylation modulates FACT binding to nucleosomes. Here,we performed dynamic atomic force microscopy and NMR analyses to clarify how the densely phosphorylated AID masks theDNA binding interface of the high-mobility-group domain (HMG). Dynamic atomic force microscopy of the nearly intact FACTrevealed that a small globule temporally appears but quickly vanishes within each mobile tail-like image, corresponding tothe HMG-containing IDR. The lifespan of the globule increases upon phosphorylation. NMR analysis indicated that phosphor-ylation induces no ordered structure but increases the number of binding sites in AID to HMG with an adjacent basic segment,thereby retaining the robust electrostatic intramolecular interaction within FACT even in the presence of DNA. These data lead tothe conclusion that the inhibitory effect of nucleosome binding is ascribed to the increase in the probability of encounter betweenHMG and the phosphorylated IDR.

INTRODUCTION

In recent years data derived from bioinformatics, proteinphysical chemistry, and tertiary structure analyses, suchas x-ray crystallography and NMR, have led to the conclu-sion that numerous functional proteins contain regions ordomains that do not form well-defined orderly three-dimensional structures but rather exist as dynamic ensem-bles of interconverting flexible conformers (1–3). Theseintrinsically disordered regions (IDRs) are ubiquitouslyfound in eukaryotic proteins that play crucial roles ingene regulation within nuclei (4–8). It is known that theseIDRs are involved in functionally important molecularrecognition, where they fold into ordered conformationsthrough binding to rigid ordered protein subunits or do-mains, a process that is generally termed coupled folding(9,10). Conversely, IDRs in a number of proteins do notnecessarily undergo global disorder-to-order transitionsupon binding. Moreover, several important regulatory in-teractions involve dynamic complexes in which mainchains of IDRs continue to fluctuate without formingdefined architectures (11,12). These binding events appearto be modulated by critical posttranslational modifications

Submitted October 10, 2012, and accepted for publication April 4, 2013.6Manami Hashimoto, Noriyuki Kodera and Yasuo Tsunaka contributed

equally to this work.

*Correspondence: [email protected] or [email protected].

ac.jp

Editor: Daniel Muller.

� 2013 by the Biophysical Society

0006-3495/13/05/2222/13 $2.00

including phosphorylation, acetylation, methylation, andubiquitination (8,11,13).

Our previous work reported that high-speed atomic forcemicroscopy (HS-AFM) can simultaneously visualize eachbehavior of IDRs and rigid domains of Drosophila chro-matin transcription-facilitating protein (dFACT) on a sub-strate surface in solution (14). This method is suitable forstudying proteins involved in gene expression, such as tran-scriptional regulators and chromatin-remodeling factors.For example, FACT, classified as a chromatin remodeler,is a heterodimer complex that consists of SSRP1 andSPT16 subunits (Fig. 1). Both subunits contain several rigiddomains and IDRs that play crucial roles in chromatinremodeling and transcriptional elongation (15–19). Thus,FACT is a suitable target for investigating the mechanisticand functional aspects of IDRs in solution through HS-AFMand NMR analyses.

In a previous study on interactions between dFACT andnucleosomes, we found that dFACT initially binds nucleo-somes and/or nucleosomal DNA via a high-mobility-groupdomain (HMG) and an HMG-flanking basic ID segment(BID) of the dSSRP1 subunit, which jointly forms aDNA-binding element (Fig. 1) (20). The acidic ID segment(AID), adjacent to BID, forms intramolecular interactionswith both HMG and BID (Fig. 1). Extensive phosphoryla-tion of AID strikingly increases the number of negativecharges, thereby strengthening the intramolecular interac-tions. As a result, the binding of dFACT to nucleosomal

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FIGURE 1 A schematic drawing of the domain organization of dSPT16

and dSSRP1, the two subunits in dFACT. The fragments used in this work

are listed: AID; SB-HMG (residues 543–554), which contains a part of the

BID region colored in purple (residues 519–554); andAB-HMG.These frag-

ments jointly constitute the regulatory region of dSSRP1 binding to DNA.

The heterodimeric complex, consisting of dSPT16 (residues 401–887) and

the full-length dSSRP1, was used for HS-AFM measurements.

Phosphorylated ID region dynamics 2223

DNA is blocked, preventing the formation of rigid structures(20). The physiological significance of this control mecha-nism is highlighted by rapid chromatin transactions duringearly embryogenesis through dephosphorylation in thematernally transmitted dSSRP1 after fertilization (20). Inhumans, the dephosphorylated SSRP1, including theHMG, is essential for FACT to exhibit higher binding affin-ity with nucleosomes (19). In contrast, the yeast FACT-HMG functions as an isolated Nhp6a/b protein (15,21,22).Thus, the phosphorylation-dependent regulation is likelyto be conserved only in higher eukaryotes. Despite theimportance of AID phosphorylation, the detailed blockingmechanism remains unresolved at the atomic level.

To clarify the mechanism of intramolecular interactionbetween the AID segment and the DNA-binding elements,we investigated the detailed effects of phosphorylation bycombining NMR spectroscopy and HS-AFM. Specifically,we prepared large amounts of various fragments, whichwere labeled with stable isotopes for NMR in both the phos-phorylated and nonphosphorylated states, whereas FACTmolecules nearly intact in size were observed by HS-AFM.Thus, this study allowed us to conclude that dense phosphor-ylation causes an electrostatic reinforcement of the interac-tions between the acidic ID region and the DNA-bindingelements, and that the essence of the interactions is dynamicand transient, thus lacking specific and stable binding sitesin domains or fragments.

MATERIALS AND METHODS

Protein preparation

The dFACT used was the same as that described in detail in our previous

study (20). The fragments used in this work are schematically drawn in

Fig. 1: AID, the short basic IDR plus HMG (SB-HMG), and the acidic

and basic IDRs plus HMG (AB-HMG). These fragments were expressed

in Escherichia coli. Details of the purification of each fragment and phos-

phorylation of the AID and AB-HMG fragments by casein kinase II (CK2)

are described in the Supporting Material.

For the dynamic AFM experiments, the His-tagged Drosophila SPT16

(401–887) (dSPT16) and Drosophila SSRP1 (dSSRP1) were cloned into

a pFastBacDual plasmid. To obtain the Ser/Thr-to-Ala mutants at the

10 phosphorylation sites (10SA; S443A, S472A, S476A, T477A, S488A,

S496A, S500A, S502A, S506A, and S515A (Fig.S1 in the Supporting

Material)), site-directed mutagenesis of the dFACT proteins was performed

by the QuikChange method (Stratagene, La Jolla, CA). The nearly intact

dFACT heterodimers (His-dSPT16 (401–887) þ dSSRP1) were coex-

pressed as the fully phosphorylated form (wild–type (WT)) and a nonphos-

phorylated form (10SA) in Sf9 insect cells and purified as described

previously (20).

Dynamic AFM analysis of the dFACT heterodimer

Dynamic AFM imaging experiments of the nearly intact dFACT hetero-

dimers (Fig. 1) were carried out as described previously (14) using a lab-

oratory-built HS-AFM (23,24). Briefly, we diluted the dFACT-WT or

dFACT-10SA samples to ~2 nM with buffer A (20 mM Tris-HCl,

pH7.5, 50 mM KCl, 10 mM MgCl2, and 0.5% glycerol (vol/vol)). The

diluted samples were used within 3 h. A droplet (2 mL) of a diluted sample

was deposited on a freshly cleaved mica surface (~1 mm in diameter

and <0.05 mm in thickness), which had been glued onto a glass stage

(2 mm in diameter and 2 mm in height). After incubation for 3 min, mol-

ecules that were not attached to the mica surface were removed by rinsing

with ~20 mL of buffer A. The sample surface was not allowed to dry. Sub-

sequently, the sample stage was immersed in a liquid cell filled with

~60 mL of buffer A in which a small cantilever had been fixed. Imaging

was carried out by HS-AFM in tapping mode. Details of the HS-AFM

imaging experiments were summarized in a recent work (25). For image

analysis, we applied three filters in the following order. First, a low-pass

filter to remove spike noise; second, a flattening filter to make the xy-plane

flat; third, a line-to-line base collection filter to minimize the base height

difference between lines.

NMR spectroscopy

The backbone 1H/13C/15N resonance assignments of AID and phosphory-

lated AID fragments were performed with a standard set of triple resonance

experiments on a DMX600 spectrometer (Bruker, Billerica, MA). (BMRB

entry 11511) Details are available in the Supporting Material.

NMR titration experiments

The interactions between the isolated AID and SB-HMG fragments were

performed in two ways: one experiment used nonlabeled AID titrated

into 15N-labeled SB-HMG, and the other experiment involved titrating non-

labeled SB-HMG into 15N-labeled AID. To explore how phosphorylation of

AID influenced its interaction with binding partners, a phosphorylated AID

fragment was also used in the titration experiments. The chemical-shift

changes observed in a series of two-dimensional (2D) 1H-15N heteronuclear

single-quantum coherence (HSQC) spectra during the titration were

analyzed by numerical curve fitting to elucidate the number of effective

binding sites, n, and the dissociation constants, KD, according to a previ-

ously published procedure (26). In the calculations, we applied a global

fitting procedure that incorporates all the chemical-shift-change profiles

for the residues under study, with n and KD as global adjustables (27).

Monte Carlo simulation was applied to estimate the statistical errors for

n and KD, assuming an uncertainty for each peak position on a 1H-15N

HSQC spectrum of 0.002 ppm (1H) and 0.02 ppm (15N), respectively,

and using 64 synthetic data sets with Gaussian noise (28). Experimental de-

tails in the titration experiments between the isolated AID and SB-HMG

fragments are described in the Supporting Material.

Biophysical Journal 104(10) 2222–2234

2224 Hashimoto et al.

NMR analysis on the interaction between AB-HMGand double-stranded DNA

Wecarried outNMRspectral analysis for the 15N-labeled nonphosphorylated

andphosphorylatedAB-HMGfragments in the absence andpresence ofDNA

(Fig. 1). The limited solubility and the severe spectral overlap in the spectra

of AB-HMG did not permit complete assignment of the resonances arising

from the backbone nuclei. Only a limited number of resonances were as-

signed for AB-HMG. We used double-stranded DNA (dsDNA) with the

sequence 50-d(CGCGATATCGCG)2-30. The reaction mixture including the

dsDNA was passed through a NAP-10 column to exchange the buffer to

50 mM Tris-HCl, pH 6.6, the buffer used for the NMR experiments.

RESULTS

Molecular features of phosphorylated andnonphosphorylated dFACT heterodimersvisualized by HS-AFM

The wild-type dFACT (dFACT-WT) is spontaneously phos-phorylated in Sf9 insect cells and is expressed as the fullyphosphorylated form at 10 phosphorylation sites. Incontrast, a dFACT mutant containing Ser/Thr-Ala substitu-tions at each of these sites, dFACT-10SA, is expressed asthe nonphosphorylated form, as described previously (20).The molecular features of dFACT-WT (the phosphorylatedform) and dFACT-10SA (the nonphosphorylated form)attached to mica surfaces in the buffer solution were directlyvisualized by HS-AFM (Fig. 2, A and B, respectively; alsosee Movies S1 and S2). At first glance, it appears thatboth molecules consist of a large globular domain and along tail region. The large globular domain appears to bewell adsorbed onto the mica surface, and its position doesnot change significantly over time. In contrast, the longtail region exhibits rapid fluctuations because of thermalagitations, indicating that this region has less affinity tothe mica surface. These molecular features are consistentwith the results of our previous study (14). Consideringour previous data (14), we can assign the large globulardomain (termed GD1, see Fig. 2 C) to the structured domainconsisting of dSPT16 (401�887) and dSSRP1 (1�404)(Fig. 1). The long tail region can be assigned to the IDRconnected to the HMG (Fig. 1).

With careful observations, we noticed that the IDR hastwo small globular domains, which were connected withIDR1–2 and IDR2–3 (Fig. 2 C). One of the small globulardomains (GD3 (see Fig. 2 C)) was consistently observed atthe end of the IDR and its appearance is not substantiallydifferent between the two constructs. In our previous study,this small globular domain was not observed, although thesample was prepared using the E. coli expression system.Furthermore, the amino acid sequence indicates no tertiarystructure at the end of the IDR. Taken together, we postulatethat unknown posttranslational modifications would inducea globular domain at the end of the IDR.

On the other hand, the other small globular domain (GD2

(Fig. 2 C)) appears temporally around the middle of the IDR


but shortly vanishes. In other words, we observed that theheight of GD2 changes over time. In addition, the figure ofGD2 appears to be retained for a longer period in dFACT-WT than in dFACT-10SA, whereas the length of the IDRappears to be slightly shorter in dFACT-WT than indFACT-10SA. Notably, the distance between GD1 andGD2 appears to be much shorter in dFACT-WT; however,the distance between GD2 and GD3 is similar in both con-structs. Similar tendencies were observed for most of theother protein molecules examined in each construct.

Quantitative analysis of distinct HS-AFM imagesbetween phosphorylated and nonphosphorylateddFACT

To quantitatively evaluate minute variations of the above-mentioned molecular features, we performed image anal-ysis. Because the IDRs are highly flexible and can adoptsimilar molecular features at an arbitrary time, successiveAFM images must be analyzed without skipping imagesto maintain arbitrariness of data sampling. To identifyIDRs that contribute to changes in molecular features, astraightforward approach is required for contour lengthanalysis of IDRs. However, considering the spatiotemporalresolution of the HS-AFM, it is currently not possible toapply contour-length analysis to successive AFM imageswithout skipping images. As an alternative, we created asimple schematic representation of the molecular featuresof dFACT observed by HS-AFM (Fig. 2 C). In this sche-matic, the heights of the three globular domains (H1–H3)and the end-to-end distances between the two globulardomains (D1–2, D2–3, and D1–3) were obtained from a singleAFM image. This approach allowed us to analyze almost allAFM images, even when the AFM images had poor signal/noise (S/N) ratios. The end-to-end distance can be used toadequately evaluate the length of a biological polymer ona surface, as described previously (29), and our data there-fore support our interpretation. The analysis was performedon three typical molecules for the phosphorylated (dFACT-WT) and nonphosphorylated (dFACT-10SA) constructs, andthe results are summarized in Table 1. We believe that thethree molecules selected lie within the population as a wholeof each construct, because most of the molecules observeddisplayed similar molecular features. For example, wecan see four molecules of dFACT-10SA simultaneously(Movie S3), all of which appear to have similar molecularfeatures. Note that the parameters described herein are notlikely to have reached the most probable values, becausewe analyzed 4586–5770 images of only three moleculesfor each construct. However, we believe that these parame-ters will be convincing for our conclusions because theyrepresent well the molecular features of each construct.The details are described below.

The distributions of H1 appeared to be similar for bothconstructs, and the mean height was 3.9 nm (Fig. 3, A

FIGURE 2 Typical HS-AFM images showing the molecular features of dFACT-WT (A) and dFACT-10SA (B). These HS-AFM images were clipped from

the movie files (see Movie S1 for dFACT-WTand Movie S2 for dFACT-10SA). Every image was taken at 67.08 ms/frame (~15 frames/s). The time from the

beginning of the clip is indicated at the upper left of each image. Scanning area, 100� 100 nm2 with 80� 80 pixel; Z-scale, 4.0 nm. The observed molecular

features of dFACTare schematized according to the definitions in C. These schematics were drawn freehand by tracing the AFM images by visual estimation.

(C) Top-view (upper) and side-view (lower) schematics simply represent the characteristics of dFACT observed by HS-AFM. Gray-colored ellipses and

black-colored thick solid lines represent the GDs and the IDRs, respectively. The symbols used for image analysis are also depicted. For every image,

we selected three points (P1–P3) (green dots) representing the peak heights (Hi) in their respective GDs. Using these points, a distance from the highest point

of one GD to that of another was determined. For example, the distance from P1 of GD1 to P2 of GD2 was expressed as D1–2. In some images, the appearance

of GD2 or GD3 was unclear; there was no point with distinct heights around the area usually seen. In this case, P2 and P3 with the highest height around the

middle and the end of the IDR were selected.


and D, and Table 1), indicating no significant difference inthe GD1 for the two constructs. This height is the same asthat obtained in our previous study (14). Although the distri-butions of H3 appeared to differ, the mean height was1.6 nm, with similar standard deviations for both constructs(Fig. 3, C and F, and Table 1), indicating that the GD3

domain may not differ between the two constructs. Incontrast, distributions for H2 were notably higher fordFACT-WT than for dFACT-10SA (Fig. 3, B and E). Twopeaks could be seen in the H2 distribution of dFACT-10SA around 1.1 nm and 1.7 nm (Fig. 3 E). The peak around1.1 nm was higher and wider than that around 1.7 nm, andthe ratio of the areas of the two peaks was ~2:1 (low/high)(Fig. 3 E). Because we observed at least two physical states

in GD2 (i.e., a lower-height and a higher-height state), thisresult indicates that the GD2 of dFACT-10SA tends towarda lower-height state than a higher one. On the other hand,the H2 distribution of dFACT-WT appears to be a combina-tion of a large peak around 1.6 nm and a small peak around1.1 nm because a small shoulder around 1.1 nm could beseen in the distribution (Fig. 3 B). The area of the largepeak was more than twice as wide as that of the smallpeak, indicating that the GD2 of dFACT-WT tends towarda higher-height state than a lower one. Consequently, theseresults suggest that the GD2 of dFACT-WT resides in thehigher-height state much longer than does that of dFACT-10SA. Considering the two height values seen in the H2

distributions (i.e., 1.1 nm for the lower-height state and


TABLE 1 Summary of the AFM image analysis

Parameter dFACT-WT dFACT-10SA

H1 3.9 5 0.3 nm 3.9 5 0.3 nm

H3 1.6 5 0.3 nm 1.6 5 0.4 nm

D1–2 15 5 5 nm 17 5 5 nm

D2–3 10 5 3 nm 11 5 4 nm

D1–3 21 5 5 nm 23 5 7 nm

D1–2/(D1–2 þ D2–3) 0.59 5 0.11 0.62 5 0.11

Number of molecules analyzed 3 3

Number of frames analyzed 5770 4586

Heights and distances are shown by mean 5 SD. Units of areas are arbi-

trary. The accuracy of the HS-AFM measurement for the z-direction (i.e.,

background noise) was 0.15 nm. The standard deviations of the Gaussian

fits were therefore restricted to >0.15 nm. Note that we did not apply

Gaussian fittings to the H2 distributions because the results obtained had

no statistical relevance.


1.6–1.7 nm for the higher-height state), the height of theHMG on one side corresponded to ~1 nm (30) and the heightof the IDR was determined to be 0.4–0.6 nm. These datacollectively suggest that the lower-height state (H2 of~1.1 nm) and higher-height state (H2 of 1.6–1.7 nm) repre-sent the HMG alone and the HMG associated with the IDR,respectively.

The distributions of D2–3 were similar between dFACT-WT and dFACT-10SA (Fig. 3, H and K), and the meanlength of D2–3 showed almost the same values (Table 1).The length of IDR2–3 did not change in response to phos-phorylation. Meanwhile, the mean length of D1–2 ofdFACT-WT (15 5 5 nm) was shorter by ~2 nm than thatof dFACT-10SA (175 5 nm) (Fig. 3, G and J, and Table 1).Therefore, the shorter state of IDR1–2 within dFACT-WT(the phosphorylated form) is generated by the interactionbetween HMG and IDR1–2, which contains the phosphory-lated AID, but not that between HMG and IDR2–3. Themean length of D1–3 of dFACT-WT (21 5 5 nm) was alsoshorter by ~2 nm than that of dFACT-10SA (23 5 7 nm)(Fig. 3, I and L, Table1). This difference can be ascribedto the difference in the D1-2 for the two constructs becausewe observed no difference greater than 2 nm in D2-3.

To estimate the mean position of the GD2 in the long taildomain, the ratios of D1–2/(D1–2 þ D2–3) for dFACT-WT(0.59 5 0.11) and dFACT-10SA (0.62 5 0.11) were deter-mined (Fig. 4), indicating that phosphorylation of IDR1–2

shifts the mean position of GD2 to GD1 but not to GD3.Combined with the results in Fig. 3, these results demon-strate that the height change for GD2 and the length changesfor D1–2 in dFACT-WT result from the interaction betweenthe HMG and IDR1–2, but not between the HMG andIDR2–3. This observation is in good agreement with our pre-vious model (20).

To summarize the HS-AFM data, GD2 temporally ap-pears and then vanishes in both constructs. However, it isimportant that the lifetime of the higher-height state indFACT-WT is much longer than that in dFACT-10SA. Inaddition, dFACT-WT seems to have a shorter IDR than


dFACT-10SA. These data indicate that the higher-heightstate of GD2 originates from the interaction between theHMG and IDR1–2, which contains the phosphorylated AID(Fig. 1). Analysis by HS-AFM is highly suitable for directlyobserving the dynamic behavior of large protein complexeson the surface of a substrate in solution. The findingsdescribed here should be analyzed by NMR, which canprovide detailed local and atomic information on protein-protein interactions.

Structural characterization of the phosphorylatedAID fragment

The fragments used in the NMR analysis are schematicallydrawn in Fig. 1. These fragments were expressed in E. coliand were phosphorylated by CK2 in vitro (Fig. S2). Thelimited signal dispersion of the isolated AID fragment inthe 2D 1H-15N HSQC spectrum is typical of a disorderedprotein (Fig. 5 A). The 15N{1H} heteronuclear nuclear Over-hauser effect (NOE) values for most residues of the AIDwere negative, indicating that the fragment is essentiallyin a disordered state. Residues around 453 showed positiveheteronuclear NOE values. This region is rich in hydropho-bic amino acids and is therefore likely to have restrictedlocal backbone motions; although the motional propertiesdo not seem functionally relevant (Fig. 5 C). CK2 success-fully phosphorylated all nine canonical phosphoacceptorserines within the AID, as revealed from high-field NMRchemical shift changes in both the 1H and 15N dimensions(Fig. 5 B); T477 was not phosphorylated in this samplepreparation, presumably due to the lower activity of CK2toward Thr residues (31). The phosphorylation of AID didnot induce any structural changes, because the increase inthe resonance dispersion in the 2D 1H-15N HSQC spectrum(data not shown) was negligible. The 15N{1H} heteronu-clear NOE profile for the phosphorylated AID (pAID) alsoindicated that it is in a disordered state (Fig. 5 C). Nonethe-less, the heteronuclear NOE values of pAID did show anoverall small but significant increase in the size of theNOE values when compared with the data for the nonphos-phorylated form (Fig. 5 C). Presumably, electrostatic repul-sion among phosphate groups may induce some fragmentstiffness.

Interactions between the isolated AIDand SB-HMG fragments

To elucidate the binding modes between isolated fragments,both the AID and pAID were titrated to the 15N-labeledSB-HMG (Fig. 6, A and B), which contains a short basicIDR and the HMG (Fig. 1). The normalized chemical-shiftchanges observed in the 2D 1H-15N HSQC spectra of15N-labeled SB-HMG in the titration (Fig. 6, A and B) areplotted versus the residue numbers (Fig. 6, C and D). Thecomparison between the two profiles showed that AID and

FIGURE 3 Height distributions of three GDs and distributions of distance between two GDs of dFACT. Height distributions (H1–H3) are shown for

dFACT-WT (A–C) and dFACT-10SA (D–F). Distance distributions (D1–2, D2–3 and D1–3) are shown for dFACT-WT (G–I) and dFACT-10SA (J–L). Green

lines represent single-Gaussian fitting. Note that we did not apply Gaussian fittings to the H2 distributions because the results obtained had no statistical

relevance. A summary of the analysis is presented in Table 1. These results were obtained from analysis applied to three molecules for each construct.


pAID share similar binding sites on SB-HMG, irrespectiveof phosphorylation. Although they do have contact withthe DNA binding surface of the HMG, both fragments pri-marily bind to the BID, because the residues in the BIDshowed greater chemical shifts than those in the HMG(Fig. 6, C and D). Furthermore, we found in our previouswork that the BID is essential for intramolecular interactionwith the AID (20). The magnitudes of the chemical shiftchanges by the addition of pAID are overall greater thanthose observed for AID binding, indicating that pAID islikely to have higher affinity to SB-HMG. That is, theincrease in the population of the bound form of SB-HMGwith pAID causes greater spectral changes relative to thefree-state spectrum (27).

The numerical fittings to the chemical shift changesagainst the molar ratios of AID or pAID to SB-HMG

showed that phosphorylation alters their binding modes toSB-HMG (Fig. 6, E and F) (26,32). To determine the KD

and n values from the experiments by using 15N-labeledSB-HMG, we included in our calculation 11 residues forAID and 30 residues for pAID, showing that the significantchanges deviated more than the mean þ 1 SD (Fig. 6, C andD, red bars). The number of SB-HMG-binding sites on AIDwas estimated at n ¼ 2.05 0.1 with a dissociation constantof KD ¼ 9.7 5 0.3 mM, assuming that all the binding siteson AID have equivalent affinities for SB-HMG. In the caseof SB-HMG binding to pAID, the number of binding siteswas estimated at n ¼ 3.9 5 0.1 with KD ¼ 1.01 50.02 mM. This indicates that phosphorylation of AID dou-bles its number of SB-HMG-binding sites in addition toenhancing its affinity for SB-HMG. In agreement with thesedata from NMR, isothermal calorimetry (ITC) experiments


FIGURE 4 Plots showing the ratio of D1–2/(D1–2 þ D2–3), which indicate

the mean positions of GD2 in the tail regions of dFACT-WT (A) and dFACT-

10SA (B). Green lines represent single-Gaussian fitting. A summary of the

analysis is presented in Table 1. These results were obtained from analysis

applied to three molecules for each construct.


using the SB-HMG and AID or pAID showed that the num-ber of binding sites on AID approximately doubles, depend-ing upon phosphorylation (data not shown).

The SB-HMG titration to 15N-labeled AID or pAID wasalso examined. The spectral changes in the 2D 1H-15NHSQC spectra for AID and pAID are shown in Fig. 7, Aand B, respectively. The 20 residues in AID showed signif-icant spectral changes (chemical-shift changes deviating bymore than the mean þ 1 SD of the entire data) upon bindingto SB-HMG. It is thus likely that these residues are involvedin binding to SB-HMG (Fig. 7 E). The residues are classifiedinto two groups according to their affinity (nine high-affinityresidues (Fig. 7 C, left, and Fig. 7 E, blue bars) and 11 low-affinity residues (Fig. 7 C, right, and Fig. 7 E, red bars)).The global fitting calculation, using the chemical shift pro-files for the nine high-affinity residues gave a KD value of3.7 5 0.1 mM, whereas the KD value for the 11 low-affinityresidues was 435 1 mM. It should be noted that one-to-onebinding was assumed in these calculations. The residuesshowed linear spectral changes according to SB-HMG con-centration (Fig. 7 A), indicating that the high- and low-affin-ity residues do not form individual binding sites. If residueswith different affinities form individual binding sites, thechemical-shift-change profiles should become nonunidirec-tional (33). Therefore, the binding surfaces on AID forSB-HMG may contain both high- and low-affinity residues.In fact, the residues of 15N-labeled SB-HMG (Fig. 6 C, redbars) showed a slightly lower affinity for AID (KD ¼ 9.750.3 mM) than did the high-affinity residues in AID (KD ¼3.7 5 0.1 mM). These results imply that the high- and


low-affinity residues of AID do not interact with SB-HMGin an individual manner but form multiple complexes in adynamic equilibrium (i.e., a dynamic complex) (34).

The titration experiments using 15N-labled pAID showedthat pAID could accommodate up to four SB-HMG mole-cules (Fig. 7, B and D), which is consistent with the resultfrom the analysis using 15N-labeled SB-HMG (n ¼ 3.9 50.1). Some residues in pAID showed nonunidirectionalspectral changes in the titration of SB-HMG (Fig. 7 B).These residues are localized in the C-terminal half, whichis rich in phosphoserine and acidic residues (Fig. 7 F, greenbars). Similar to the nonlinear spectral changes in multipleligands binding to the protein (27,35), it is likely thatmultiple SB-HMGs bind to the phosphor-Ser-rich regionof pAID. The nonunidirectional spectral changes may there-fore be caused by the expansion of the phosphor-Ser-rich re-gion in response to the multiple bindings of SB-HMG.

Taken together, these data allowed us to conclude that non-phosphorylated AID contains binding surfaces that canaccommodate up to two SB-HMGmolecules, whereas phos-phorylation of AID expands the surface by a factor of 2. Thebinding surfaces expanded by phosphorylation should in-crease the intramolecular encounter probability betweenthe AID and BID-HMG segments in the dSSRP1 subunit.

Changes of intramolecular interactions betweenAID and BID-HMG in response to phosphorylationand dsDNA binding

To explore how AID forms intramolecular interactions withBID-HMG, we compared the 2D 1H-15N HSQC spectra ofan AB-HMG fragment and the isolated AID (Fig. S3).The limited solubility (<0.1 mM) and the longer IDR ofAB-HMG hampered assignment of the resonances of thebackbone nuclei. Only a limited number of signals for ratherisolated Ser residues were assigned based on the close prox-imity (i.e., similar chemical shifts) of the resonances for theAB-HMG construct to those assigned in the spectra of iso-lated AID or SB-HMG fragments (Fig. S3).

The significant spectral differences between AB-HMGand the isolated AID demonstrated that part of the AIDsegment binds to the other part of the AB-HMG intramolec-ularly (Fig. S3 A). The spectral difference between the twofragments remained in the presence of double-strandedDNA (dsDNA) (Fig. S3 B). This observation shows thatthe AID segment in the AB-HMG fragment still contactsother parts of the fragment without being released to behaveas an isolated fragment, which would give NMR signals thatmatch more closely to those for the isolated AID fragment.

The spectral difference between the pAB-HMG and theisolated pAID was apparent, indicating that the pAIDsegment in the pAB-HMG fragment has intramolecular con-tacts with other parts of the fragment (Fig. S3 C). The pres-ence of the dsDNA did not completely reverse the spectraldifference (Fig. S3 D), indicating that it does not block

FIGURE 5 NMR spectral characterization of AID fragments. (A) Backbone resonance assignments of 15N-labeled AID in the 2D 1H-15N HSQC spectrum.

(B) Spectral comparison in a region of the 2D 1H-15N HSQC spectra for nonphosphorylated (red) and phosphorylated AID (blue). The backbone amide sig-

nals of the phosphorylated Ser shows an upfield shift in both the 1H and 15N dimensions. (C) Comparison of the 15N{1H} heteronuclear NOE profiles between

nonphosphorylated (red) and phosphorylated (blue) AID fragments.



FIGURE 6 NMR titration experiments using the15N-labeled SB-HMG fragment. (A) Chemical-

shift changes in the titration of nonphosphorylated

AID fragment (AID) to 15N-labeled SB-HMG;

AID/SB-HMG molar ratios were 0.0 (black),

0.3 (red), 0.7 (blue), 1.0 (orange), 1.5 (pink),

2.0 (green), and 3.0 (yellow). (B) The correspond-

ing spectral changes observed for phosphorylated

AID (pAID); pAID/SB-HMG ratios were

0.00 (black), 0.07 (red), 0.21 (blue), 0.49 (orange),

0.70 (pink), and 1.04 (green). (C and D)

Histograms of the chemical-shift differences in

SB-HMG upon binding with AID (C) and pAID

(D). Chemical-shift differences are plotted against

residue numbers of SB-HMG. Red bars indicate

that the chemical-shift differences are over the

average plus one standard deviation. Short bars in

cyan and purple along the x axis represent prolines

and the residues for which assignment information

was missing. Chemical-shift changes are shown for

the representative residues in SB-HMG in the titra-

tions with AID (E) and pAID (F). The numerically

determined dissociation constant, KD, and number

of binding sites, n, for the AID and pAID titration

experiments were KD ¼ 9.75 0.3 mM, n ¼ 2.0 5

0.1 and KD ¼ 1.01 5 0.02 mM, n ¼ 3.9 5 0.1,

respectively. Values for KD and n were determined

by global fitting using the residues marked by red

bars in C and D (see Supporting Material).


the intramolecular contacts of the pAID segment with partsof the pAB-HMG fragment.

The absence of a complete set of resonance assignmentsfor the AB-HMG and pAB-HMG fragments allows only alimited analysis of the intramolecular interactions betweenthe AID and BID-HMG segments. Despite the experimentallimits of this study, the spectral comparison shows that intra-molecular interactions between the segments happen irre-spective of phosphorylation, and that these interactionsremain even in the presence of dsDNA; however, the modesof binding seem to change.

The intramolecular interactions were monitored using alimited number of assigned HMG signals (Fig. S4). In addi-tion to the limited solubility of the AB-HMG fragment,intense signals from the long unstructured part in the AB-HMG prohibited observation of the signals from the HMG.

Despite the limited residues monitored, these NMR datacollectively indicate that, irrespective of the phosphorylation


state, the residues in HMG and BID retain contact withthe AID segment, as suggested by the spectral differencein the HMGs in the AB-HMG and pAB-HMG fragmentsrelative to the isolated HMG (Fig. S4). In the presence ofdsDNA, the intramolecular contacts of HMG with the AIDsegment appear to change according to the phosphorylationstate of the AID segment, as evinced by the spectral com-parisons for residues A606, K621, and G553 (Fig. S4).The phosphorylation-dependent spectral changes for theHMG induced by the addition of dsDNA suggest that phos-phorylation of the AID segment alters its interaction withthe HMG box and thus changes the HMG-mediated bindingto dsDNA.

DISCUSSION AND CONCLUSIONS

The combination of HS-AFM and NMR analyses has shownthat phosphorylation of multiple serine residues in the AID

FIGURE 7 Spectral changes on 15N-labeled AID

and pAID in the titration with the unlabeled

SB-HMG. (A) Spectral changes induced by the

titration of AID; SB-HMG/AID ratios were

0.0 (black), 0.1 (green), 0.3 (red), 0.7 (cyan),

1.0 (yellow), 1.5 (purple), 2.0 (blue), and

3.0 (pink). (B) Spectral changes by pAID;

SB-HMG/pAID ratios were 0.0 (black), 0.3 (green),

1.0 (red), 2.0 (cyan), 3.0 (yellow), 5.0 (purple),

7.0 (blue), and 9.0 (pink). (C) Chemical-shift-

change profiles for representative residues in AID,

showing the high (left) and low (right) affinities to

SB-HMG. (D) The 15N chemical-shift changes

observed for representative residues in pAID

according to the titration. Spectral changes deter-

mined that the maximal binding number of

SB-HMG to pAID is 4. (E and F) Histograms of

the chemical-shift differences in AID (E) and

pAID (F) upon binding with SB-HMG. Chemical-

shift differences are plotted against residue

numbers for AID and pAID. Residues with a

mean þ 1 SD are colored. Residues in AID with

blue bars have higher affinity (KD ¼ 3.7 5

0.1 mM) than those with red bars (KD ¼ 43 5

1 mM). The residues in pAID with green bars

showed kinked titration traces, as in B. Short

down bars in cyan and purple along the x axis indi-

cate prolines and unassigned residues, respectively.

Short down bars in red indicate phosphorylated Ser

residues. Short down orange bars are residues

whose signals were incompletely traced due to

severe signal overlap during the titration.


modulates the intramolecular interactions between the AIDand the DNA-binding element, which consists of BID andHMG. Our data collectively provide mechanistic insightsinto dynamic interactions between AID and the DNA-bind-ing elements in FACT, as follows.

1. AID makes intramolecular contact with the DNA-bind-ing elements. Phosphorylation of AID expands the bind-ing epitope to the DNA-binding element by a factor of 2when compared with the nonphosphorylated region.

2. The intramolecular contact of the nonphosphorylatedAID with the DNA-binding element should be relatively

weak, as demonstrated by the limited spectral changes inresonances from the isolated AID (Fig. S3). This isconsistent with our HS-AFM observation that the non-phosphorylated FACT retains mostly extended IDRs(Fig. 3). Thus, the intramolecular contact between thenonphosphorylated AID and the DNA-binding elementremains dynamic and transient.

3. Phosphorylation of AID reinforces its intramolecularinteraction with the DNA-binding element, as shownby the larger spectral changes for the resonances repre-senting the phosphorylated serines in the AID whencompared with the same resonances in the isolated


FIGURE 8 Summary of the mechanism underly-

ing phosphorylation-dependent inhibition with

respect to nucleosomal DNA. The intramolecular

contact of nonphosphorylated AID with BID-

HMG is nonspecific and dynamic. This situation

allows BID-HMG to transiently bindwith the nucle-

osomal DNA. In contrast, dense phosphorylation

expands the AID region by elevating repulsive

forces. This results in an increase of the probability

of encounter between BID-HMG and pAID, thereby

forming a more robust complex between BID-HMG

and pAID. AID and BID are symbolically repre-

sented by the red and purple strings, respectively.

Red and purple spheres indicate negative and posi-

tive net charges, respectively. The HMG is denoted

by the cyan L-shaped structure. Red open circles

labeled P indicate phosphorylation sites.


pAID (Fig. S3). The phosphorylated AID retained intra-molecular contacts with the DNA-binding elements inthe presence of DNA (Fig. S3), thus indicating that thisAID element impairs FACT binding to DNA. In the pres-ence of DNA, several resonances representing residueswithin the HMG showed chemical-shift changes, despitethe fact that these residues retain contact with phosphor-ylated AID (Fig. S4). Our HS-AFM observation revealeddynamic behavior between folding and unfolding of theglobular domain, which is putatively formed by pAIDand BID-HMG (Fig. 3). Taken together, FACT involvesdynamic and transient intramolecular interactions, evenin the phosphorylated state.

The structure of the dSSRP1-HMG was similar to theyeast Nhp6a protein structure isolated from SSRP1 (30).The DNA-binding surface on the dSSRP1-HMG structurewas similar to the DNA-binding surface in Nhp6A. TheNMR structure of the Nhp6A-DNA complex revealed acharacteristic L-shaped HMG fold, which contacts theminor groove of DNA, whereas an extended N-terminalBID region interacts with the adjacent major groove (36).The DNA-binding interface contains numerous conservedlysine and arginine residues that participate in electrostaticinteractions with the phosphate backbone. Although somehydrophobic stacking/wedge interactions are formedbetween a number of Nhp6A residues and DNA bases, theinteractions of Nhp6Awith DNA are predominantly electro-static. Our NMR data indicate that DNA and AID occupyalmost the same electrostatic interaction interfaces on theHMG and BID regions regardless of the phosphorylationstate of AID.

An apparently similar mechanism was proposed for theautoinhibition of DNA binding by phosphorylation of a tran-scription factor, Ets-1 (37,38). The affinity of Ets-1 to DNAis allosterically regulated by the flexibility of the unstruc-tured serine-rich region and the inhibitory module, whichis adjacent to the ETS domain responsible for DNA binding


(38). Previous NMR experiments demonstrated that phos-phorylation at several sites within the serine-rich regiongradually shifts the equilibrium more to the rigid-inactiveform, which is stabilized by stronger intramolecular interac-tions with both the inhibitory module and the DNA-bindingdomain (39). However, this inhibitory effect of Ets-1 isconsiderably different from our previous results, whichshowed no additive reduction of DNA-binding inhibitionin response to the number of phosphorylated Ser/Thr muta-tions of dFACT (20). In the FACT mutant containing theSSRC motif, AID, BID, and HMG (Fig. 1), Ser/Thr-to-Ala mutations at two phosphorylation sites showed negli-gible inhibition of DNA binding, whereas mutations atfour and six sites dramatically decreased the inhibitoryeffect (as shown in Fig. 3 C of Tsunaka et al. (20)). In otherwords, the inhibitory effect appears to be ultrasensitive, butnot linear.

Our HS-AFM data provide, to our knowledge, a novelstructural view, where the dynamic behavior of the FACT-IDR is drastically altered in response to phosphorylation.The NMR data showed that dense phosphorylation of theAID region increases binding sites with the DNA-bindingelements containing the HMG. Presumably, the phosphory-lation expands the AID region by elevating the repulsiveforces, thus inducing a stronger interaction without proteinfolding. Moreover, the increase in binding sites caused bythe phosphorylation of residues can be ascribed to a dy-namic equilibrium among multiple binding states but not asimple two-state equilibrium, thereby blocking DNA bind-ing (Fig. 8). This notion relates to the concept of encounterprobability and differs to the mechanism observed for Ets-1.

This dynamic binding mode is rather similar to that inthe electrostatic model proposed for the interaction be-tween Cdc4 and Sic1 (34,40); the SCF ubiquitin ligasesubunit Cdc4 interacts with the cyclin-dependent kinase in-hibitor Sic1 in a dense phosphorylation-dependent manner(34). Multiple phosphorylation in the N-terminal IDRsof Sic1 leads to equilibrium engagement by an


interchange between phosphorylation sites (34). The de-gree of phosphorylation fine-tunes the complex formationof Sic1 with Cdc4 via long-range electrostatic interactions,ensuring ultrasensitivity of the Sic1-Cdc4 interactioncaused by a net charge reversal (40). In agreement withthis ultrasensitive change, our previous data showed thatsimultaneous mutations at four or six phosphorylation sitesdrastically enhance the binding ability of FACT to nucleo-somal DNA (as shown in Fig. 3 C of Tsunaka et al. (20)).It is thus likely that the inhibitory mechanism of thephosphorylated FACT is essentially based on the electro-static model.

Furthermore, a notable feature of this regulation mecha-nism is that the cooperative action of the tandemly linkedAID and BID regions directs nucleosomal DNA bindingthrough their interaction with the HMG in FACT. Thesetandemly linked IDRs may enhance the probability ofencounter between the DNA-binding elements and the phos-phorylated IDR in the inhibitory mechanism (Fig. 8). In fact,our HS-AFM analyses indicate that the lifetime of the glob-ular domain in phosphorylated dFACT is much longer thanthat of the nonphosphorylated form.

Many atomic structures of protein complexes provide ageneral view that domain and/or subunit contacts take placeon well-ordered surfaces that complement each other. How-ever, functionally important complexes frequently involvedynamic, or transient, properties, which are derived fromunstructured IDRs even upon complex formation. It isvery likely that such flexibilities of protein complexesshould change in response to posttranslational modifica-tions, thereby regulating physiological functions. Thus, thecombination of NMR and HS-AFM analysis used in thisstudy potentially can be applied to many other protein com-plexes in which IDRs play important roles.

SUPPORTING MATERIAL

Four figures, three movies, supporting materials, and methods are avail-

able at http://www.biophysj.org/biophysj/supplemental/S0006-3495(13)

00433-5.

This work was supported by Grants-in-Aid for Scientific Research on Inno-

vative Areas (Research in a Proposed Research Area) (Ministry of Educa-

tion, Culture, Sports, Science and Technology KAKENHI Grant No.

21113002, 21121006, and 23107724); a Grant-in-Aid for Japanese Society

for the Promotion of Science Fellows (JSPS KAKENHI Grant No.

07J01196); the Mitsubishi Foundation; Japan Science and Technology

(JST) Precursory Research for Embryonic Science and Technology; and

JST Development of Systems and Technology for Advanced Measurement

and Analysis.

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Phosphorylation-Coupled Intramolecular Dynamics of